Dissection and Immunohistochemistry of the Drosophila Adult Leg to Detect Changes at the Neuromuscular Junction for an Identified Motor Neuron

The procedures for preparing working solutions used in conjunction with this protocol are described in Table 1.

Table 1. Solutions for performing immunocytochemistry of the adult Drosophila leg.

1. Initial fixation of tissues

1. Select approximately 10 flies for each genotype and age. Anesthetize on a fly pad under carbon dioxide (Figure 1A, B).
   NOTE: Begin with more flies than necessary to ensure a large enough sample size after dissection.
2. Using a paintbrush, transfer flies to cold methanol in a glass well or dish for approximately 30 seconds to 1 minute (Figure 1C). The methanol solubilizes the cuticular hydrocarbons and flies can now be submerged rather than float in aqueous solutions.
3. With forceps, carefully transfer flies to PBS. Rinse 3x in ice-cold PBS to remove excess methanol and keep flies in PBS on ice until dissection and fixation. At this point, dissect flies and fix within the shortest time possible (<30 minutes).
4. To isolate legs, transfer flies to a silicone elastomer dissection dish filled with cold PBS, remove the metathoracic legs at the coxa using two pairs of #5 Dumont forceps or cut the legs with Vanna scissors (Figure 1D). Transfer the legs to a well in a plastic 24 well plate filled with 1 mL of PBS and keep the plate on ice until all legs are removed and transferred to wells.
   NOTE: Each well can hold at least 20 legs.
5. Replace the PBS solution with 1 mL of FA solution and rotate on a nutator for 30 minutes (Figure 1E). The nutator setting should be set at a medium speed (17 rpm). Ensure the legs are completely submerged in FA solution during this time for adequate fixation.
6. To remove FA solution, wash in 1 mL of PBS 3x quickly followed by an additional 3 washes for 5 minutes each in 1 mL of PBS. Hold tissues in 1 mL of PBS on ice prior to, and during the dissection steps described below.

2. Removing the Leg Cuticle and Antibody Staining

1. Removing the leg cuticle
   1. The dissecting forceps are critical for success. Introduce slight parallel bends in both prongs at the end of #5 super fine forceps to provide a bevel that allows the cuticle to be grabbed superficially rather than be poked, which can ruin the tissue (Figure 1F, G).
      NOTE: Prongs bent in parallel should still make contact with each other throughout the length of the prong when closed (Figure 2).
   2. Transfer legs to a silicone elastomer dish in PBS for dissection. Orient a leg so that the anterior side is facing up (see Figure 3 for leg anatomy and orientation information). Using one pair of forceps, hold the tibia segment against the silicone elastomer dish. Using the other forceps held bevel side down, grab a piece of cuticle on the distal end of the femur and pull in the proximal direction toward the trochanter.
   3. Keep methodically removing cuticle until the naked muscle is visible throughout the proximal end of the femur (Figure 1F, G, H).
      NOTE: Only make superficial contact with the legs using the beveled side of the forceps to avoid pulling muscle.
   4. Once all legs are dissected, replace PBS with FA to post-fix legs for 30 minutes with shaking on a nutator at medium speed (Figure 1I). Wash samples in 1 mL of PBS for 3 times quick and then 3 times for 5 minutes each in PBT (Figure 1J).
      NOTE: If staining with monoclonal antibody NC82 (anti-bruchpilot) to label active zones, post-fix for 20 minutes as this antigen is sensitive to longer fixations.

Figure 2. Modified forceps used for dissecting adult legs. (A) The ends of the forceps are bent and then flattened at the bottom (arrow) creating a bevel by filing on a sharpening stone. (B) In contrast, the prongs of unmodified forceps are not bent. Scale bar = 1 mm Please click here to view a larger version of this figure.

2. Antibody staining
   1. To block tissues for antibody staining, replace 1 mL of PBT with blocking solution consisting of 1 mL of 5% NGS diluted in PBT. Incubate dissected legs for 4 hours at room-temperature or overnight at 4 °C while rocking on a nutator at medium speed (17 rpm). During all incubations, cover wells with sealing tape in addition to the plastic cover (Figure 1J).
      NOTE: 24 well plates are used for immunocytochemistry rather than 1.5 mL or 2 mL microcentrifuge because previous attempts to use microcentrifuge tubes resulted in broken legs and damaged tissue.
   2. Remove blocking solution and add 300 mL of primary antibodies diluted in fresh blocking solution. The small volume of antibody used should be sufficient to cover the tissues. Re-seal wells with laboratory sealing tape and plastic lid and incubate overnight at 4 °C with shaking on a nutator at medium speed (Figure 1J). The working antibody reagent information and concentrations used in for these studies are described in Table 2.
   3. Wash primary antibodies in 1 mL of PBT for 3 times briefly and then 3 times for 15 minutes each (Figure 1J).
      NOTE: Diluted primary antibodies can be saved and reused if stored at 4 °C for up to 2 weeks.
   4. Block tissues again in 1 mL of 5% NGS for at least 2 hours at room temperature or overnight at 4 C (Figure 1J).
   5. Remove the 5% NGS blocking solution and add 300 µL of the appropriate diluted fluorescent conjugated secondary antibodies. Additionally, add 1:2000 dilution of fluorescence-conjugated phalloidin to label muscle (Figure 1J).
   6. For secondary antibody incubations, seal wells with laboratory sealing tape and lid. Also, wrap plates in aluminum foil to protect fluorophores from light. Incubate for 6-8 hours at room-temperature or overnight at 4 °C.
   7. Wash secondary antibodies and phalloidin as described in step 2.2.3 above. Cover plates with aluminum foil in between washes to protect fluorophores from light.

3. Mounting Legs

1. Transfer legs to a slide using forceps and orient anterior side up. Cover the legs with mounting media (Figure 1K, L).
   NOTE: The dissected legs can be aspirated with a P1000 pipette tip if the bottom bore is widened by cutting with a razor blade.
2. Add clay spacers to a 22×22 mm² coverslip (#1.5 thickness) by scraping the coverslip corners across a small ball of modeling clay. Each corner should have a small amount of clay 1-2 mm thick (Figure 1K).
3. To cover the slide, add the coverslip with the clay spacers facing towards the slide and carefully push on the corners until the coverslip just touches the surface of the femur.
4. To prevent evaporation, seal the edges of the coverslip with nail polish and let dry in a dark place (about 10 minutes) before storing at 4 °C until imaging.

4. Imaging

1. Image by confocal microscope (Figure 1M). Include a transmitted light channel to better assess the quality of the dissection and samples with visibly disrupted muscle fibers in the area of interest should be discarded.
   1. Begin imaging z-stacks with 20x magnification with 2x zoom and a total image depth of ~ 40 mm corresponding to the thickness of the femur. For fluorescent signal detection, capture images at resolutions consistent with Nyquist sampling (we are using 1024 x 1024 pixels with a dwell setting of 8-10 μs/pixel). The signal intensity should be in the linear range which is achieved by adjusting the high voltage gain settings. Once the gain settings are set for a series of samples within an experiment, they should not be altered so that signal intensities can be compared between samples.
2. For imaging synaptic boutons and other subcellular structure, capture confocal images at ≥60x magnification. The detector settings should be in the linear range, while the pixel density, dwell settings and z-depth should be similar for images captured at lower magnification (step 4.1).